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Abstract

Sensory and cognitive impairments have been documented in diabetic humans and
animals, but the pathophysiology of diabetes in the central nervous system is
poorly understood. Because a high glucose level disrupts gap junctional
communication in various cell types and astrocytes are extensively coupled by
gap junctions to form large syncytia, the influence of experimental diabetes on
gap junction channel-mediated dye transfer was assessed in astrocytes in tissue
culture and in brain slices from diabetic rats. Astrocytes grown in
15–25 mmol/l glucose had a slow-onset, poorly reversible decrement in
gap junctional communication compared with those grown in 5.5 mmol/l glucose.
Astrocytes in brain slices from adult STZ (streptozotocin)-treated rats at
20–24 weeks after the onset of diabetes also exhibited reduced dye
transfer. In cultured astrocytes grown in high glucose, increased oxidative
stress preceded the decrement in dye transfer by several days, and gap
junctional impairment was prevented, but not rescued, after its manifestation by
compounds that can block or reduce oxidative stress. In sharp contrast with
these findings, chaperone molecules known to facilitate protein folding could
prevent and rescue gap junctional impairment, even in the presence of elevated
glucose level and oxidative stress. Immunostaining of Cx (connexin) 43 and 30,
but not Cx26, was altered by growth in high glucose. Disruption of astrocytic
trafficking of metabolites and signalling molecules may alter interactions among
astrocytes, neurons and endothelial cells and contribute to changes in brain
function in diabetes. Involvement of the microvasculature may contribute to
diabetic complications in the brain, the cardiovascular system and other
organs.

INTRODUCTION

Many diverse, progressive and severe complications of diabetes are well established
and are linked to chronically high glucose levels in conjunction with insulin
deficiency (Type 1 diabetes) or insulin resistance (Type 2 diabetes). The complex,
multifactorial pathobiology of diabetes (Brownlee,
2005) includes non-specific glycation reactions of glucose, increased
sorbitol production and osmotic stress, oxidative stress due to generation of ROS
(reactive oxygen species)/RNS (reactive nitrogen species), depletion of endogenous
antioxidants, enhanced lipid peroxidation, metabolic changes, altered hormonal
responses, cardiovascular disease, kidney damage, poor wound healing, and cataract
formation. Overall, the impact of diabetes on the central nervous system is
generally considered to be mild or modest compared with involvement of peripheral
organs and peripheral neuropathies (Little et al.,
2007), which have severe consequences for both the quality and duration
of life of diabetic patients.

Our interest in the involvement of astrocytes in diabetic complications of the
central nervous system arose from reports of impaired gap junctional communication
in hyperglycaemic vascular smooth muscle, endothelial cells and retinal pericytes
(Inoguchi et al., 1995; Stalmans and Himpens, 1997; Kuroki et al., 1998; Oku et al., 2001; Sato et al.,
2002; Li et al., 2003). We
recently found that astrocytes in the inferior colliculus, an auditory pathway
structure with the highest metabolic rate in brain, are highly coupled by gap
junctions (Ball et al., 2007) and are involved
in selective syncytial ‘trafficking’ of energy and redox
metabolites (Gandhi et al., 2009a), including
lactate (Gandhi et al., 2009b). (Note: in the
present study, metabolite ‘trafficking’ refers to transfer
among cells of small molecules involved in metabolism, energetics and signalling by
processes driven mainly by concentration gradients. Trafficking of small molecules
involves diffusion and transporters, and differs from protein
‘trafficking’. Fluorescent dyes are used as surrogate markers
to visualize and quantify movements of small molecules among cells.) We hypothesized
that diabetes may cause ‘silent’ changes affecting astrocytic
communication and metabolite trafficking via gap junctions may alter interactions
among astrocytes, neurons and endothelial cells (i.e. the neurovascular unit),
thereby contributing to the slow, progressive brain dysfunction in diabetes. The
present study, therefore, examined the effects of experimental diabetes on
astrocytic gap junctional transport using two model systems, the STZ-diabetic rat
and cultured astrocytes grown in medium containing very high glucose concentrations.

Studies in rat models of diabetes show that plasma and brain glucose levels increase
on average by approx. 3-fold, with mean values in brain rising from 2.2
μmol/g in controls to 6.9 μmol/g in diabetic animals (Table 1). In STZ-diabetic rats, the increases
in plasma and brain glucose content occur within 2 days after STZ treatment and
persist for months at levels similar to those in spontaneously diabetic rats (Table 1). The rise in brain glucose
concentration with an increase in plasma glucose level is the expected consequence
of concentration gradient-driven transport of glucose across the
blood–brain barrier. Under steady state conditions in normal rats infused
with various concentrations of glucose, the brain glucose level is approx. 20% that
in plasma in the normo- and hyper-glycaemic range; in contrast, the brain plasma
glucose distribution ratio falls during hypoglycaemia when glucose supply does not
match demand (Dienel et al., 1991, 1997; Holden et
al., 1991). Thus brain glucose level rises when plasma glucose
concentration increases, and in diabetic rats the brain: plasma glucose distribution
ratio is even higher, approx. 50% greater than in control rats, and the elevated
ratio is not normalized by acute insulin treatment to reduce plasma glucose level to
the normal range (Hofer and Lanier, 1991a,
1991b). Corresponding studies of brain
glucose level in human diabetics are sparse, but one NMR study reported a 1.5-fold
increase in the level of glucose relative to creatine in diabetic brain and
calculated a net rise in brain glucose level of approx. 2 mmol/l (Table 1). Routine commercially available tissue
culture media contain glucose concentrations as high as 25 mmol/l glucose, which
approximates to the level of glucose in the plasma of diabetic animals and exceeds
the normal and diabetic rat brain glucose level by approx. 10- and 3-fold
respectively (Table 1). Astrocytes grown in
‘high’-glucose media would be exposed to the myriad of
well-established consequences of severe, chronic hyperglycaemia, and the
pathophysiological consequences of neuronal and Schwann cell culture in high-glucose
media have been recently emphasized by Kleman et al.
(2008) and Mìinea et al.
(2002). In the present study, astrocytes chronically exposed to elevated
glucose levels in vivo and in vitro were used as
models of experimental diabetes. We report that intercellular gap junction-mediated
communication among astrocytes is markedly reduced in cultured cerebral cortical
astrocytes and in slices of inferior colliculus from STZ-treated rats, and that
pharmacological intervention can protect against or restore this impairment.

Astrocyte culture

Cultured astrocytes were prepared by small modifications of established
procedures (Hertz et al., 1998). Briefly,
astrocytes were harvested from the cerebral cortex of 1-day-old albino
Wistar–Hanover rats (Taconic Farms, Germantown, NY, U.S.A.) and grown
in T-75 culture flasks with DMEM containing 5.5 mmol/l glucose, 10% (v/v) FBS,
50 IU (international units) of penicillin and 50 μg/ml of
streptomycin at 37°C in humidified air containing 5% CO2.
l-LME (0.1 mmol/l), a lysosomotrophic agent that selectively
destroys mononuclear cells including microglia, was also included in the culture
medium, and the cultures were shaken by hand twice per week to remove microglia.
When confluent, the cells were trypsinized, seeded on to polylysine-coated glass
coverslips and grown to confluence in a medium containing amphotericin B (2.5
μg/ml). Then differentiation was induced by supplementing the culture
medium with 0.25 mmol/l dBcAMP. The next day, cells were maintained in a medium
containing 0.25 mmol/l dBcAMP and 5.5, 15 or 25 mmol/l glucose for up to 4
weeks. Purity of cultures was based on the expression of the astrocyte marker,
glial fibrillary acidic protein, which was expressed in >90% of the
cells.

STZ-induced diabetes

Male Sprague–Dawley rats (200–300 g; Harlan, Indianapolis,
IN, U.S.A.) were fasted overnight, injected intraperitoneally with STZ (65 mg/kg
body weight in 33 mmol/l citrate-buffered saline, pH 4.5); controls received the
same volume of citrate-buffered saline (Romanovsky et al., 2006). Tail blood samples were taken for glucose
determination from overnight-fasted animals on the day before STZ injection and
on days 3, 8 and 13 thereafter; rats were categorized as normoglycaemic or
hyperglycaemic, using a cut-off value of >6.9 mmol/l to define
hyperglycaemia based on the day 3 fasting blood glucose level. All animal use
procedures were in strict accordance with the NIH Guide for Care and Use of
Laboratory Animals and were approved by the local Animal Care and Use
Committee.

Brain slice preparation

At 20–24 weeks after induction of STZ-diabetes, the diabetic and
age-matched, vehicle-injected control rats were deeply anaesthetized with
halothane and decapitated and their brains were quickly removed and chilled by
immersion in an oxygenated (i.e. bubbled with O2/CO2,
95:5), ice-cold aCSF (artificial cerebrospinal fluid) solution (concentrations
in mmol/l: 26 NaHCO3, 10 glucose, 124 NaCl, 2.8 KCl, 2.0
MgSO4, 1.25 NaH2PO4 and 2.0 CaCl2,
pH 7.3) and 248 sucrose, and 250 μm-thick slices were cut using a
Leica (Heidelberg, Germany) VT 1000S tissue slicer; inferior colliculus slices
were incubated in oxygenated aCSF containing sucrose for 30 min at
35°C and then for 1 h at 22°C (Moyer and Brown, 1998). Slices of inferior colliculus were
transferred to an open bath perfusion chamber (Warner Instruments, Hamden, CT,
U.S.A.) on the microscope stage. Then the slices of inferior colliculus and the
cultured astrocytes were perfused (1 ml/min) with aCSF that was continuously
bubbled with O2/CO2 (95:5) and contained 26 mmol/l
NaHCO3 (pH 7.3) and 10 mmol/l glucose at approx.
21–22°C.

Gap junction dye transfer assays

Two procedures were used to insert a membrane-impermeant dye into astrocytes to
assay gap junctional communication, scrape-loading (el-Fouly et al., 1987; Giaume et al., 1991) and diffusion into a single cell impaled with a
micropipette (Ball et al., 2007; Gandhi et al., 2009a). Scrape loading is
commonly used for dye transfer assays because it is a simple procedure, but the
procedure destroys cells at the scrape site and releases their contents to the
medium; it requires medium changes and washing of cells after dye loading, and
it cannot be used in tissue slices in which astrocytes have formed their
syncytia during normal brain development involving interactions of astrocytes
with neurons and the vasculature. Microinjection of cells is technically more
difficult, but offers more control for dye loading, and it can be used in brain
slices. Fluorescent compounds used to assay dye transfer were LYVS (4% or 62
mmol/l), 4% LYVS plus 4% LYCH, Alexa Fluor® 350 (5 mmol/l) and 6-NBDG
(20 mmol/l). 6-NBDG is a non-metabolizable fluorescent analogue of glucose that
is a substrate for glucose transporters. In the brain slice assays of 6-NBDG gap
junctional transfer, 10 μmol/l of cytochalasin B, a glucose transport
inhibitor (Speizer et al. 1985), was
included to minimize efflux of 6-NBDG from cells via glucose transporters. In
these assays, an excess amount of pyruvate (10 mmol/l) was added to the
perfusate as an oxidative fuel to compensate for blockade of glucose transport.

Scrape loading

The procedure of Giaume et al. (1991)
was used for scrape-load assays, as follows. Glass coverslips containing
astrocytes were transferred to sterile 35 mm culture dishes; coverslips were
washed once and incubated in ionic-buffered solution containing
(concentrations in mmol/l) 130 NaCl, 2.8 KCl, 1 CaCl2, 2
MgCl2 and 10 Hepes (pH 7.2) for 1 min, washed again and
incubated in calcium-free medium for 1 min. The medium was replaced with a
calcium-free medium containing 0.5 mg/ml LYVS and 1 mmol/l
rhodamine–dextran (a non-permeant macromolecule used to label the
scrape-loaded cells); a 2–3 cm scrape was made with a razor
blade, and dye transfer was allowed for 2 min. Giaume et al. (1991) stated that calcium was omitted
from the medium during scrape-loading because 1 mmol/l Ca2+
blocks Lucifer Yellow transfer; Mg2+ was still included in the
scrape-load medium. A recent study showed that incubation of cultured
astrocytes in the nominal absence of extracellular bivalent cations
(Ca2+ and Mg2+) opens channels (hemichannels or
pannexin channels) that allow rapid, widespread entry of Lucifer Yellow into
astrocytes (Ye et al., 2003); the
presence of Mg2+ during the scrape-loading would prevent opening
of these channels. Cells were washed twice with calcium containing
ionic-buffered solution and dye-labelled area determined at 8 min after
scraping by image analysis using MetaVue software. Line scans were used to
evaluate the change in Lucifer Yellow fluorescent intensity as a function of
distance from the scrape site. The dye-labelled area was determined in three
regions of each scrape; regions were imaged, and gap junctional dye transfer
was calculated as the difference between the areas labelled by LYVS (gap
junction permeable) and rhodamine–dextran (labels the dye-loaded
cells); the mean value for each triplicate determination was used as the
area labelled in that scrape-load assay.

Dye transfer was also assayed in astrocytes cultured in media containing 5.5
or 25 mmol/l glucose for up to 21 days in the presence or absence of
compounds to reduce ROS/RNS levels or that facilitate protein folding.
MnTBAP (50 μmol/l) is a superoxide dimutase mimetic that is a
scavenger of ROS (Kowluru and Abbas,
2003) and l-NAME (1 mmol/l), a competitive inhibitor of NOS
(nitric oxide synthase) that impairs formation of nitric oxide and RNS
(Hink et al., 2001). Chemical
chaperones (Brown et al., 1996; Welch and Brown, 1996; Özcan et al., 2006), 4-PBA
(1 mmol/l), glycerol (25 mmol/l), TUDCA (25 mmol/l) and TMAO (100 mmol/l)
were added to the culture media at the time intervals indicated. Tunicamycin
(100 ng/ml, 16 h), an inhibitor of N-linked glycosylation that causes ER
(endoplasmic reticulum) stress, served as the positive control for ER
stress; butyrate (1 mmol/l) was a control for 4-PBA. After cultured
astrocytes were treated for the time intervals indicated, LYVS transfer was
assayed by impaling a single astrocyte with a micropipette, allowing the dye
to diffuse for 2 min, and then the dye-labelled area was measured using
MetaVue software.

Oxidative stress assays

Astrocytes were grown in 5.5 or 25 mmol/l glucose in the presence or absence of
inhibitors or chaperones, and ‘oxidative stress’ was
assayed with DCF-DA or carboxy-DCF-DA, compounds that are cell membrane
permeable, cleaved by intracellular esterases and, after oxidation by various
reactive compounds, become fluorescent dichlorofluorescein (DCF) or carboxy-DCF
(Tampo et al., 2003; Cruthirds et al., 2005 and references cited
in these studies). At indicated days in culture, DCF-DA (10 μmol/l)
was added to the culture medium (that had been changed 24 h earlier) or to a
fresh medium containing the inhibitors plus 30 μmol/l DCF-DA. Cells
were returned to the CO2 incubator for 30 min at 37°C,
then washed with perfusion solution, and DCF fluorescence intensity was measured
with the Nikon E600 microscope (×40 objective) and MetaVue software.
Ten field-of-view images were collected per coverslip, and analysed by
thresholding to include the pixels with the highest 30% or highest 2%
fluorescence intensity; the 30% threshold value excluded background fluorescence
and included the cell bodies plus ‘hot spots’, whereas the
2% threshold included only the highest-intensity ‘hot
spots’. Slices of inferior colliculus from diabetic rats were
incubated for 30 min in 10 μmol/l carboxy-DCF-DA and fluorescence
intensity assayed as described above.

Cx immunostaining

Cultured astrocytes on coverslips were fixed with 2% (w/v) paraformaldehyde in
0.1 M PBS for 10 min, washed three times with PTX (0.1 M PBS containing 0.3%
Triton X), blocked in 10% (v/v) goat serum in PTX for 30 min, and incubated with
rabbit polyclonal primary antibodies (diluted in 10% goat serum in PTX as
follows: Cx43, 1:250 to a final concentration of 1 μg/ml; Cx30,
1:250, to 1 μg/ml; Cx26, 1:25, to 5 μg/ml) for 2 h at room
temperature (approx. 21–22°C) and then overnight at
4°C. The manufacturer's recommended levels for use in
frozen sections were 1–5 μg/ml for Cx43 and Cx30
antibodies and 10–20 μg/ml for Cx26; in the present study,
the dilution of Cx43 was the same as that used by Ye et al. (2003). The next day, samples were warmed to
room temperature, washed with PTX (three 5 min washes), incubated with goat
anti-rabbit secondary antibodies conjugated to Texas Red (diluted 1:500 in 10%
goat serum in PTX) for 1 h at room temperature, given three 5-min washes with
PTX and stored at 4°C in PBS. Immunostained Cx protein includes
intracellular punctate or vesicle-like structures (probably ER, Golgi apparatus
and cytoplasmic vesicles; see Wolff et al.,
1998 and references cited therein), that were prominent in the images
of immunoreactive Cxs under the fixation and immunoassay conditions used in the
present study (see the Results section). The area of this punctate or vesicular
immunoreactive material was measured by image analysis of composite images of
z-stacks using the maximum projection setting with a Nikon E600 microscope with
confocal attachments and a ×60 water immersion objective (NA 1.00)
and MetaVue software. The minimal fluorescence intensity threshold value was set
to only include prominent punctate or vesicle-like structures, and integrated
morphometric analysis was used to measure their total area in each of the
16–36 cells per Cx group that were derived from two independent
cultures.

Statistics

Comparisons between two groups of independent samples were made with two-tailed,
unpaired t tests. Comparisons among three or more groups of
independent samples were made with one-way ANOVA and Dunnett's test
for multiple comparisons against the same control value or the Bonferroni test
for multiple comparisons among experimental groups.
P<0.05 was considered to be statistically
significant. All statistical analyses were performed with GraphPad
Prism® software, version 5.02 (GraphPad Software, La Jolla, CA,
U.S.A.).

RESULTS

Transcellular spreading LYVS after scrape-loading of astrocytes grown for 3 weeks
in 5.5 mmol/l glucose greatly exceeded that of astrocytes grown in 25 mmol/l
glucose (Figure 1A). Line scan analysis of
Lucifer Yellow fluorescence intensity with distance showed that overall LYVS
fluorescence level was higher and dye spread extended further from the scrape
site in cells grown in the lower glucose concentration (Figure 1B). The mean LYVS intensity in the pixels closest to
the scrape site (at 0.6 μm) tended to be higher
(P=0.084) in the low-glucose
cultures and it reached a peak (~2600 fluorescence units) at
26–29 μm from the scrape, whereas the high-glucose
cultures had a much lower mean maximal value (~1700 fluorescence
units; P<0.001) and a broader peak that was closer
(7.7–20 μm) to the scrape (Figure 1B). At 90–100 μm from the scrape the
fluorescence intensities in the low glucose group were still 22% higher
(P<0.001) than in the higher glucose group.
These findings suggest that the lower dye levels and reduced dye spread in
severely hyperglycaemic astrocytes are not due to differential release of
Lucifer Yellow from cells during the scrape load procedure via Cx
‘hemichannels’ or pannexin channels. If these channels
were preferentially open to the medium in either group of cells, extensive dye
labelling would be expected to increase markedly throughout the culture, not
just adjacent to the scrape site, as observed (Figures 1A and 1B); this labelling would be readily detected by
visual observation because the LYVS causes prominent labelling of nuclei.

Growth of astrocytes in high glucose reduces gap junctional
communication

The net area labelled by LYVS was calculated by subtracting the area of the
scrape-filled cells (i.e. area labelled by the gap junction-impermeant tracer,
rhodamine–dextran) from that labelled by LYVS. Astrocytes grown in
low glucose had a 4-fold higher dye-labelled area than those grown in high
glucose (Figure 1C). Blockade of gap
junctions with octanol reduced the LYVS-labelled area (Figure 1C) to the level of that labelled by
rhodamine–dextran (results not shown).

Prolonged exposure to high glucose is required to reduce dye transfer

Glucose concentration in the culture medium did not affect astrocyte viability,
and astrocytes grown in 5.5, 15 and 25 mmol/l glucose had similar cell densities
on days 3, 14 and 21 (results not shown), as illustrated in Figure 2 for representative cultures grown for 2 weeks in
5.5 mmol/l (Figure 2A) or 25 mmol/l (Figure 2B) glucose. However, when gap
junctional communication was assayed by impaling a single astrocyte with a
micropipette and diffusing Lucifer Yellow into one cell, dye spreading from the
impaled cell was much greater in cells grown for 3 weeks in 5.5 mmol/l glucose
(Figure 2C) compared with those grown
in 25 mmol/l glucose (Figure 2D),
confirming the results of scrape-load assays (Figure 1).

Time in culture did not affect the area labelled by Lucifer Yellow in astrocytes
grown in low glucose, but those grown in high glucose had a progressive decrease
in gap junctional communication (Figure
2E). Impaired LYVS transfer had a slow onset, requiring approx.
3–5 days exposure to 15 or 25 mmol/l glucose before a statistically
significant decrement was detectable. The time courses and maximal inhibition
for cells grown in 15 and 25 mmol/l glucose were similar; the maximal decrement
in gap junctional communication was relatively stable at approx. 50% of that in
the low-glucose cultures during the interval from 7 to 21 days (Figure 2E).

Diffusion of a smaller fluorescent dye, Alexa Fluor® 350, among
astrocytes was stable with time in the low-glucose cultures, and it also
exhibited a progressive fall in labelled area in the high-glucose cultures
(Figure 2F). There was a 5-day delay
before Alexa Fluor® 350-labelled area was reduced by high glucose,
and the 50% decrement was stable between 7 and 21 days. Thus the two dyes had
similar lag times, temporal profiles and maximal reduction of labelled area,
suggesting that reduced dye transfer may not be simply due to partial
constriction of the gap junctional channel to block the passage of larger
molecules (Alexa Fluor® 350 has a molecular mass of 311 Da after
hydrolysis of the succinimidyl ester by water compared with 536 Da for the
ionized form of LYVS). Note that Alexa Fluor® 350 does label a
greater area than the LYVS in the low-glucose cultures (e.g.
P<0.01 on day 1; compare Figures 2E and 2F); this is probably due mainly to its high
fluorescence quantum yield (the concentration of Alexa Fluor® 350 in
the micropipette was 5 mmol/l compared with 62 mmol/l for LYVS), and perhaps
also to its smaller size.

Dye transfer deficit is not restored by subsequent glycaemic control

When astrocytes were grown in 15 or 25 mmol/l glucose for 2 weeks and then
transferred to 5.5 mmol/l glucose for an additional 2 weeks, LYVS spreading via
gap junctions did not recover. Dye transfer remained subnormal after either 7 or
14 days in the low-glucose media (Figure
3), indicating that the acquired decrement in gap junctional
communication could not be reversed within 2 weeks by simply reducing the
glucose concentration in the culture medium.

Glycaemic control does not reverse the deficit previously acquired
during growth in hyperglycaemic conditions

Oxidative stress precedes impairment of gap junctional communication

Because diabetes is associated with oxidative stress (Brownlee, 2005), DCF fluorescence was assayed at intervals
after exposure of astrocytes to the high-glucose medium to assess changes in the
levels of ROS and RNS. Increased DCF fluorescence was detectable after 1 day of
exposure to high glucose, with a progressive rise with time in culture (Figure 4). Elevated ROS/RNS production
preceded impairment of gap junctional communication that became evident only
after 3–5 days of exposure to very high glucose (compare Figures 2E, 2F and ​and44).

Oxidative stress is detectable after 1 day of severe hyperglycaemia
and remains elevated

Focal oxidative stress

The high-glucose cultures had focal ‘hot spots’ of
greater DCF fluorescence intensities (quantified by thresholding the highest
2% fluorescence values; see the Materials and methods section) that averaged
approximately twice those of the overall mean intensities in the
high-glucose cultures (thresholded at 30%) at all time points (Figure 4). The hot spots in the
hyperglycaemic cultures also had quite large S.D. values. In contrast, hot
spots in the control cultures grown in 5.5 mmol/l glucose had fluorescence
intensities closer to the overall mean intensities (Figure 4).

Magnitude of oxidative stress is variable in culture
batches

In a replicate experiment (results not shown) in which astrocytes were grown
in high or low glucose for 1, 3, 7 or 14 days
(n=3–5
coverslips per group with 9–11 regions of interest assayed per
coverslip for a total of 44–70 regions per group) the cells were
incubated for 30 min in 30 μmol/l DCF-DA, and at each time point,
DCF fluorescence was statistically significantly higher in cells grown in 25
mmol/l glucose compared with those in 5.5 mmol/l glucose
(P<0.001). However, the mean DCF fluorescence
intensities in the high-glucose cultures were more similar to each other at
each time point, differing somewhat from the data set in Figure 4. The high glucose/low glucose
ratio of DCF values in the replicate cultures was 1.86, 2.84, 2.04 and 2.67
at 1, 3, 7 and 14 days for threshold at 30%, and 2.52, 4.02, 2.42, and 4.78
respectively for the ‘hot spots’ quantified by
thresholding at 2%. Thus the high-glucose cultures have higher DCF
fluorescence than those grown in low glucose, but replicate assays can
differ in magnitude. For unidentified reasons, some astrocytes grown in high
glucose exhibited low DCF fluorescence at 14 days, but they still had
reduced trafficking of LYVS. Unfortunately, the oxidative stress and gap
junctional transfer assays do not permit longitudinal studies on the same
cells; the properties of the cells before the assay are unknown.

The body weight of STZ-diabetic rats during the 13–20-week interval
after onset of diabetes averaged 53±2%
(n=4) of age-matched,
vehicle-injected controls
(n=4), and at 20 weeks, the
respective body weights were 268±36 and 471±19 g. At the
time of assay for dye transfer and oxidative stress, the arterial plasma glucose
level in diabetic rats was elevated 4.1-fold compared with controls
(33.1±5.2 and 8.0±0.9 mmol/l respectively;
P<0.001), whereas arterial plasma lactate content
was unchanged (2.2±0.7 and 2.2±0.4 mmol/l respectively).
To verify that brain glucose levels were also elevated in our STZ rats, the
glucose concentration was assayed in ethanol extracts of cerebral cortex
dissected from funnel-frozen brains of two STZ rats, using previously described
methods (Dienel et al., 2007). The brain
glucose concentrations were 8.2 and 6.8 μmol/g, indicating that both
plasma and brain glucose levels in the STZ rats used in the present study were
within the range of the mean literature values tabulated in Table 1.

Dye transfer among gap junction coupled astrocytes was assayed by dye diffusion
into single astrocytes in slices of the inferior colliculus from age-matched,
vehicle-treated control and STZ-diabetic rats. Both Lucifer Yellow and 6-NBDG
had greater dye labelling in slices from control rats (Figures 5A and 5C) compared with those from STZ-treated rats
(Figures 5B and 5D). The number of
LYVS-labelled cells was 7.7-fold higher in slices from control compared with
diabetic rats (Figure 6A), whereas the area
labelled by the fluorescent glucose analogue, 6-NBDG (342 Da), was 2.2-fold
greater in control compared with diabetic rat slices (Figure 6B). DCF fluorescence in diabetic brain slices was
3.2 times that in controls (Figure 6C).
Thus gap junctional communication was reduced and oxidative stress was increased
in slices of the inferior colliculus from diabetic rats at 20–24
weeks after STZ treatment, as observed in cultured astrocytes that were exposed
to much higher glucose levels for short time intervals (compare with Figures 2 and ​and44).

ER stress impairs gap junctional communication

Tunicamycin is an inhibitor of N-acetylglucosamine
transferases known to cause ER stress by blocking the formation of
N-glycosidic protein–carbohydrate linkages and
preventing the glycosylation of newly synthesized proteins in the ER.
Astrocytes were grown for 2 weeks in low glucose and then treated with
tunicamycin for 16 h and dye transfer was assayed. The dye-labelled area was
reduced by tunicamycin to the low level observed in vehicle-treated cells
that were grown in high glucose for 2 weeks (Figure 7A), i.e. the Lucifer Yellow-labelled area was approx.
4000 μm2 under both these conditions. For comparison
with these values, astrocytes grown in low glucose for up to 3 weeks had a
Lucifer Yellow-labelled area of approx. 15000
μm2 (Figures
2 and ​and3)3) and the 2–3
week values for low-glucose cultures from Figures 2 and ​and33 are
included in Figures 7(A) and 7(B) as
reference values. However, to avoid additional multiple statistical
comparisons against the same data sets, comparisons in Figures 7(A) and 7(B) were made against the
vehicle-treated control grown in high glucose or the no treatment group in
the high-to-low glucose transfer assay.

Influence of ROS/NOS inhibitors and chemical chaperones on dye
transfer and DCF fluorescence

ROS/RNS inhibitors and chemical chaperones are protective in the
presence of high glucose

Continuous 2-week treatment of cultured astrocytes grown in high glucose with
MnTBAP, a superoxide dismutase mimetic, or with l-NAME, an
inhibitor of NOS (alone or in combination) prevented the
high-glucose-induced decrement in dye transfer (Figure 7A). Similarly, four chemically different
chaperone molecules (4-PBA, glycerol, TMAO and TUDCA) also protected against
dye transfer impairment, whereas butyrate, a control for 4-PBA, did not
(Figure 7A).

Chaperones, not ROS/RNS inhibitors, restore the acquired
deficit

Astrocytes were grown for 2 weeks in high glucose and then transferred to the
low-glucose medium containing vehicle or other compounds for 7 days. The
dye-labelled area in vehicle-treated astrocytes remained at the low level
(Figure 7B) obtained for astrocytes
previously grown in high glucose (compare with Figure 3). Inclusion of MnTBAP or l-NAME in the
low-glucose medium did not restore gap junctional trafficking (i.e. the
dye-labelled area remained low), whereas inclusion of two chaperones, 4-PBA
and TUDCA, did improve dye transfer and increased the labelled area (Figure 7B).

Oxidative stress is eliminated by transfer to low glucose

DCF fluorescence was elevated in cells grown in high glucose for 3 weeks
compared with those grown in low glucose. However, when astrocytes were
grown in high glucose for 2 weeks and then transferred to low glucose for an
additional week, the level of oxidative stress fell to that of cells
continuously grown in low glucose for 3 weeks (Figure 7C). Thus the decrement in gap junctional communication
caused by prior exposure to high glucose persists (Figure 7B, no treatment group), even though oxidative
stress is eliminated (Figure 7C) by
reducing the glucose level in the culture medium.

Effect of gap junction permeability modulators on oxidative
stress

The ROS/RNS blockers would be expected to improve gap junctional
communication by reducing or preventing the rise in DCF fluorescence in
cells grown in high glucose, whereas the chaperone molecules would not be
expected to alter glucose-induced oxidative stress. To test these
predictions, DCF fluorescence was assayed in two independent experiments
using different batches of astrocytes that were grown for 2 weeks in 25
mmol/l glucose in the continuous presence of each of the test compounds
shown in Figure 7(A)
(n=30–40
samples per group per experiment). Unfortunately, the results in the
replicate assays were variable (results not shown), and further work is
required to identify factors that influence the response of DCF fluorescence
intensity to these drug treatments.

Summary of results of pharmacological studies

Both prolonged exposure to high glucose and short-term tunicamycin treatment
impair astrocytic gap junctional communication. In high-glucose cultures,
oxidative stress is detectable on day 1 (Figure 4), whereas the fall in dye transfer becomes manifest at
approx. 3–5 days (Figures 2E and
2F). Transfer of cells from high- to low-glucose medium was
sufficient to reduce DCF fluorescence to control levels (Figure 7C), but not restore dye transfer
to normal within 2 weeks (Figure 3).
ROS/RNS inhibitors could prevent this deficit if included in the medium at
the onset of exposure to high glucose (Figure
7A), but could not improve the acquired deficit that persisted in
the low-glucose medium (Figure 7B).
Chaperone treatment could, however, restore the gap junctional deficit in
the presence of high glucose (Figure
7A), as well as in the presence (Figure
7A) or absence (Figures 7B and
7C) of elevated DCF fluorescence. Thus prolonged antecedent oxidative
stress is linked to reduced gap junctional trafficking, but reducing ROS/RNS
levels after the onset of the deficit did not restore dye transfer (Figure 3). In sharp contrast, the
persistent change in Cx function acquired by growth in high glucose was
ameliorated by four different chaperone molecules that can improve protein
folding

Immunoreactive Cx protein

All astrocytes exhibited immunostaining for Cx43, Cx30 and Cx26, with the most
intense immunoreactivity mainly in intracellular material, as illustrated for
astrocytes grown for 2 weeks in 5.5 mmol/l glucose (Figure 8, left column). Intracellular immunostained Cx
protein can include ER, Golgi apparatus and cytoplasmic vesicles (Wolff et al., 1998 and references cited
therein), and punctate, vesicle-like intracellular staining in astrocytes is
evident in other studies (e.g. Ye et al.,
2003). The area of the immunoreactive punctate structures in astrocytes
grown in high glucose was reduced to approx. 50% and 25% of that in the
low-glucose cultures respectively for Cx43 and Cx30, whereas that for Cx26 was
unaffected (Figure 8, right column). Thus
the morphological appearance of immunoreactive Cx protein is selectively
influenced by medium glucose concentration, perhaps reflecting changes in
trafficking or turnover of these proteins.

DISCUSSION

The two major findings of the present study are that chronic hyperglycaemia and
STZ-induced diabetes markedly reduce gap junctional dye transfer among
astrocytes and that the impairment of gap junctional communication can be
prevented and rescued by pharmacological treatment with compounds that reduce
oxidative stress or improve protein folding. Impaired transcellular
communication had a slow onset and, once established, it was poorly reversible
by subsequent glycaemic control. This deficit was detectable with three tracers
of different sizes and charges (Lucifer Yellow, Alexa Fluor® 350 and
6-NBDG, a non-metabolizable glucose analogue), and the scrape-loading assays
indicate that it did not arise from differential dye release by hyperglycaemic
cells via pannexin pores or Cx hemichannels (Figures 1–​3).3). Because increased DCF fluorescence
preceded the onset of the decline in gap junctional permeability by
3–5 days and ROS/RNS blockers could prevent but not rescue the
decrement (Figures 4 and ​and7),7), damage arising from oxidative stress may
be a causative factor. Acute tunicamycin treatment generates abnormal newly
synthesized proteins, causes ER stress and impairs dye transfer within 16 h
without hyperglycaemia and oxidative stress. However, results of our ongoing
studies indicate that expression of selected markers for ER stress is delayed
compared with onset of reduced gap junctional communication in hyperglycaemic
cultured astrocytes, suggesting that gap junctional impairment may be an early,
relatively selective event in the pathophysiology of diabetes.

High glucose is sufficient to impair gap junctional communication

The effects of chronic hyperglycaemia and complications of diabetes are very
complex, and relationships among pathophysiology, threshold glucose level and
cumulative exposure to elevated glucose concentrations are very difficult to
define. However, tissue culture experiments demonstrate that severe, chronic
hyperglycaemia itself is sufficient to disrupt gap junctional communication in
astrocytes in the absence of endocrine dysfunction and multiorgan interactions.
Both of our experimental model systems, cultured astrocytes and STZ-rats, have
high glucose concentration as a variable, but they differ with respect to
maximal glucose level, cumulative exposure and effects of interactions among
brain cell types and among body organs. Cumulative exposure can be expressed as
the product of glucose concentration multiplied by time, or the area under a
plot of concentration as a function of time. Different pathophysiological
processes can be expected to take place at various threshold levels of glucose
concentration. For example, glucose flux into the sorbitol pathway will
progressively increase as glucose concentration rises above normal due to the
high Km of aldose reductase for glucose
(~25 mM). The threshold concentrations and cumulative exposure
required to cause various effects of elevated glucose (e.g. non-enzymatic
glycation reactions, oxidative stress and disruption of signalling pathways) are
unknown, but these effects could be expected to increase with glucose level and
duration of exposure (Mìinea et al.,
2002). Brain glucose levels are lower than in peripheral tissues
owing to the restrictive transport properties of the blood–brain
barrier (Table 1), but diabetic patients
live with the disease for decades, facilitating cumulative CNS (central nervous
system) effects of chronic hyperglycaemia.

Growth of cultured cells under severely hyperglycaemic conditions is a
pathophysiological condition relevant to diabetes. Commercially available tissue
culture media can contain glucose concentrations ranging from 0 to 25 mmol/l
and, for example, DMEM is formulated with 0, 5.56 or 25 mmol/l glucose,
Ham's nutrient mixtures can have 7, 10 or 17.5 mmol/l glucose and
Neurobasal™ medium (Brewer et al.,
1993) contains 25 mmol/l glucose. Even a
‘low-glucose’ medium, 5–6 mmol/l glucose, is
approximately twice the normal rat brain glucose concentration (i.e. approx.
2–3 μmol/g) and is equivalent to severe diabetes in rat
brain (Table 1). Growth of astrocytes in
22 mmol/l glucose reduces both glucose and lactate oxidation by approx. 50%
compared with cells grown in 2 mmol/l glucose (Abe et al., 2006), and would be expected to predispose astrocytes
grown in high glucose to increased glycolytic metabolism and greater lactate
release. In cultured neurons, the lactate dehydrogenase isoenzyme pattern was
not altered by medium glucose level (5.5, 13.4 or 26.8 mmol/l; O’Brien et al., 2007), but Kleman et al. (2008) emphasize the negative
effects of high glucose levels on the viability of cultured neurons and neuronal
responsiveness to the AMPK (AMP-activated protein kinase) energy signalling
system. High glucose may or may not influence experimental outcome, but diabetic
complications are, nevertheless, a concern for astrocytes and neurons grown in
high glucose, and it is important to re-evaluate the results of such studies
(e.g. 20 mmol/l glucose: Ye et al.,
2001, 2003, 2009; McCoy and
Sontheimer, 2010; 25 mmol/l glucose: Sorg and Magistretti, 1991; Yu et
al., 1993; Takahashi et al.,
1995; Itoh et al., 2003; Pellerin and Magistretti, 2005, 1994; Chenal and Pellerin, 2007; and 50 mmol/l glucose: Bliss et al., 2004). Also, Methods sections
in published studies sometimes only identify the ‘generic’
culture medium (e.g. DMEM) without stating the glucose level or other key
constituents; ideally, the catalogue number of the medium should be reported so
its formulation is available. The use of normal brain tissue glucose levels for
growth of cultured cells with twice-weekly feeding schedules may also have
nutritional complexities. For example, our unpublished data (K.K. Ball, N.F.
Cruz and G.A. Dienel) indicate that astrocytes grown in 5.5 mmol/l glucose
consume most of the glucose within approx. 12–18 h, with release of
lactate to the medium; this lactate can be subsequently consumed as an oxidative
substrate, along with glutamine and other substrates in the medium. Daily
replenishment of glucose and other nutrients and removal of lactate and other
compounds released to the culture medium may be necessary to control levels of
extracellular metabolites, but total medium replacement could also negatively
affect the cells due to various ‘stresses’ associated with
removal from the incubator and medium change, e.g. shear stress to the surface
of the cells, transient loss of CO2 and buffering capacity, and
transient hypoxia and hypothermia.

Abnormal proteins and therapeutic potential

Covalent protein modification can arise from various causes known to occur in
diabetes, e.g. non-enzymatic glycation reactions and formation of advanced
glycation end-products, protein carbonylation reactions, and altered regulation
of gene expression and signalling pathways causing abnormal phosphorylation or
nitrosylation states (Bonnefont-Rousselot,
2002; Brownlee, 2005). The ability
of four different chaperone molecules that can facilitate protein folding in
other experimental systems to (i) prevent the decline in dye transfer even in
the presence of high glucose levels and oxidative stress and (ii) rescue an
established deficit (Figure 7) suggests
that changes to Cx proteins secondary to oxidative stress may cause abnormal
protein structure, folding, protein–protein interactions or protein
trafficking that may be reflected by the morphological changes in intracellular
non-junctional immunoreactive Cx 43 and 30 protein (Figure 8). Further work is required to evaluate the
contributions of these possibilities to altered non-junctional immunoreactive
material in diabetic astrocytes. Poor reversibility of gap junctional
communication after reversion to low-glucose culture media (Figures 3 and ​and7B)
7B)
underscores the importance of continuous, strict glycaemic control in diabetic
patients. The effectiveness of treatment with chaperones (Figure 7) that are already approved for human use (e.g.
4-phenylbutyrate and TUDCA; Özcan et
al., 2006) opens a therapeutic avenue to improve gap junctional
intercellular trafficking that is effective in the presence of high glucose
levels, oxidative stress and metabolic disturbances.

Involvement of different Cxs in many cell types during experimental diabetes

Dysfunction of any or all the three astrocytic Cxs (Cx43, Cx30 and Cx26), as well
as Cx-associated proteins, may contribute to the functional deficit of gap
junctional trafficking of small molecules during experimental diabetes and would
be anticipated to affect transcellular communication via channels that comprise
these Cxs in all body tissues, not just brain. Because Cx30 channels are not
permeable to LYCH (Manthey et al., 2001),
the abnormal transfer of Lucifer Yellow may involve Cx43 and Cx26 channels that
are permeant to this dye (Elfgang et al.,
1995). Cx43 may be a major ‘target’ of diabetes in
astrocytes, as well in other organs, as suggested by previous studies in other
cell types.

Gap junctional communication in a number of cell types is inhibited by growth in
high-glucose media ranging from 14 to 30 mmol/l for different intervals
(1–9 days) compared with cells grown in 5–5.5 mmol/l
glucose, and decrements have been documented in endothelial cells in the aorta
(Inoguchi et al., 1995, 2001), in the retina (Fernandes et al., 2004) and in epididymal fat pads (Sato et al., 2002; Li and Roy, 2009), in smooth muscle cells in aorta (Kuroki et al., 1998; Inoguchi et al., 2001), in pigment epithelial cells in
retina (Stalmans and Himpens, 1997; Malfait et al., 2001), and in pericytes in
retina (Li et al., 2003). These studies
have linked dye transfer deficits to altered PKC (protein kinase C) signalling,
increased phosphorylation of Cx43, reduced Cx43 mRNA and protein levels, low
Cx43 plaque counts, and increased proteosome-mediated degradation of Cx43.
Consistent with the above findings are reports that propagation of calcium waves
is inhibited in hyperglycaemic and PKC-activated retinal pigment epithelial
cells (Stalmans and Himpens, 1997), as
well as in PKC-activated astrocytes (Enkvist and
McCarthy, 1992). In STZ-diabetic rats, dye transfer is reduced in
acutely isolated pericytes in retinal microvessels after 5–18 days of
diabetes (Oku et al., 2001). Also, the
increased duration of QRS waves in electrocardiograms from STZ-diabetic rats is
associated with increased phosphorylation of Cx43 that is linked to activation
of PKC, with either unchanged or reduced Cx43 protein levels (Inoguchi et al., 2001; Lin et al., 2006, 2008; Howarth et al.,
2008). However, in coronary endothelial cells isolated from STZ-diabetic
mice, Cx40 is a critical element in loss of gap junction intercellular
communication; its levels are reduced, along with those of Cx37, but not Cx43,
and high glucose impairs capillary network formation in vitro
(Makino et al., 2008). Together, the
above findings indicate that gap junctional communication is abnormal in many
organ systems exposed to prolonged hyperglycaemia and experimental diabetes,
with tissue- and organ-specific effects. The brain is generally considered to be
affected by diabetes to a lesser extent than peripheral organs, but gap
junctional trafficking among astrocytes, retinal cells and endothelial cells is
markedly reduced.

Roles of gap junctions in pathophysiology of diabetes and
Alzheimer's disease

Gap junction-coupled astrocytes are involved in integration of neurotransmission,
energetics and blood flow at a local level, and impaired syncytial communication
by means of cytoplasmic signalling, redox and energy-related molecules can
contribute to brain dysfunction. For example, lack of
Ins(1,4,5)P3 signalling arising from mutations
in Cx26 in non-neuronal support cells in the cochlea is sufficient to cause
deafness (Beltramello et al., 2005),
indicating that a sensory loss associated with neurons can arise from
dysfunction of other cell types whose roles are required for processing of
sensory information. Gap junctions have important roles in regulation of
vascular function (Figueroa and Duling,
2009), and the brain's vasculature is surrounded by astrocytic
endfeet that are extensively coupled with each other by gap junctions that
facilitate long-distance dye transfer along the vasculature when dye is diffused
into a single astrocyte (Ball et al.,
2007). Thus it is likely that signals among cells within the
‘neurovascular unit’ that comprises neurons, astrocytes
and endothelial cells would be disrupted by impairment of gap junctional
communication between astrocytic processes and their endfeet. As discussed
above, hyperglycaemia induces abnormalities in Cx proteins and signalling in
astrocytes, endothelial cells in different tissues, vascular smooth muscle
cells, and retinal pericytes. Microvascular pathology is common to diabetes and
Alzheimer's disease (Luchsinger and
Gustafson, 2009; Sonnen et al.,
2009), STZ-diabetic rats exhibit increased levels of amyloid
β-peptide and phosphorylated tau protein (Li et al., 2007), hyperglycaemia exacerbates
pathophysiological changes and cognitive decline in pre-symptomatic
Alzheimer's mice (Burdo et al.,
2009), and aged Alzheimer model mice have altered astrocytic networks
(Peters et al., 2009). Taken
together, these findings suggest that impairment of astrocytic gap junctional
trafficking may contribute to the pathology of the microvasculature in brain
and, ultimately, to sensory and cognitive decline in diabetes and
Alzheimer's disease. In addition, involvement of abnormalities in gap
junctional communication in vascular endothelial and smooth muscle cells and
cardiac cells may underlie or contribute to complications of diabetes in the
cardiovascular system and other organs.

Footnotes

This work was supported by NIH (National
Institutes of Health) [grant numbers NS36728,
NS47546 and DK081936]; Alzheimer
Foundation [IIRG-06-26022]; and
the University of Arkansas for Medical Sciences
Department of Physiology and Biophysics, Graduate School,
and Research Council. The content of the present work is solely the
responsibility of the authors and does not necessarily represent the official
views of the National Institute of Neurological Disorders and Stroke, the
National Institute of Diabetes and Digestive and Kidney Diseases, or the
NIH.